Biochemistry 2008, 47, 7617–7625 13C
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Relaxation Studies of the DNA Target Sequence for HhaI Methyltransferase Reveal Unique Motional Properties† Zahra Shajani‡,3 and Gabriele Varani*,‡,§ Departments of Chemistry and Biochemistry, UniVersity of Washington, Seattle, Washington 98195-1700 ReceiVed October 11, 2007; ReVised Manuscript ReceiVed April 28, 2008
ABSTRACT:
The goal of this work was to examine if sequence-dependent conformational flexibility in DNA plays a role in base extrusion, a common conformational change induced by many DNA-modifying enzymes. We studied the dynamics of the double-stranded DNA target of the HhaI methyltransferase by recording an extensive set of 13C NMR relaxation parameters. We observe that the cytidine furanose rings experience fast (picosecond to nanosecond) motions that are not present in other nucleotides; the methylation site experiences particularly high mobility. We also observe that the bases of guanosine and cytidine residues within the HhaI recognition sequence GCGC experience motions on a much slower (1-100 µs) time scale. We compare these observations with previous solution and solid-state NMR studies of the EcoRI nuclease target sequence, and solid-state NMR studies of a similar HhaI target construct. While an increased mobility of cytidine furanose rings compared to those of other nucleotides is observed for both sequences, the slower motions are only observed in the HhaI target DNA. We propose that this inherent flexibility lowers the energetic barriers that must occur when the DNA binds to the HhaI methyltransferase and for extrusion of the cytidine prior to its methylation.
Methylation of DNA is an important step in a host of biological processes, ranging from gene silencing to Xchromosome inactivation (1). The methyltransferase modification enzymes attach methyl groups to either the N4 or C5 position within the cytosine or the N6 position within adenine. The chemical addition of a methyl group to DNA bases is carried out by restriction-modification systems comprising a methyltransferase (Mtase) which methylates the DNA at specific recognition sites and an endonuclease (2). Among the best studied methylation systems is HhaI1 (Haemophilius hemolyticus), which is part of a defense mechanism in these bacteria that protects the cell form invasive foreign DNA. The HhaI system is comprised of the two canonical enzymes, the methyltransferase and endonuclease, where the former is expressed first and methylates the native DNA. The endonuclease is subsequently expressed and cleaves all nonmethylated foreign DNA. Both proteins recognize the sequence 5′-GVCGC-3′ (the arrow indicates cleavage, and the underlined cytosine base is the methylation target). The covalent addition of a methyl group to the C5 position of the target cytosine inhibits †
Supported by NIH Grant RO1 EB003152. * To whom correspondence should be addressed. E-mail: varani@ chem.washington.edu. Telephone: (206) 543-7113. Fax: (206) 685-8665. ‡ Department of Chemistry. 3 Present address. The Scripps Research Institute, Department of Molecular Biology, 10550 N. Torrey Pines, La Jolla, CA 92037. § Department of Biochemistry. 1 Abbreviations: AdoMet, S-adenosyl-L-methionine; CSA, chemical shielding anisotropy; HhaI, Haemophilius hemolyticus; HSQC, heteronuclear single-quantum correlation spectroscopy; T1, longitudinal relaxation time; T1F, rotating frame relaxation time; NOE, nuclear Overhauser effect.
FIGURE 1: Structure of DNA-HhaI methyltransferase complex (Protein Data Bank entry 1FJX). (a) DNA-protein complex, in which the C6 base flips out ∼170° in the presence of the methyltransferase. (b) Target DNA with the methyltransferase protein removed for clarity.
endonucleolytic cleavage, protecting the bacterial DNA from its own endonuclease. The methyltransferase DNA target forms a classical double helix, providing little access to the cytidine position to be methylated. However, the crystal structure of the HhaI Mtase complexed with its cognate double-stranded DNA (dsDNA) and methyl donor cofactor S-adenosyl-L-methionine (AdoMet) showed that the cytosine becomes exohelical in complex with the methylase and becomes surrounded by residues within the enzyme active site (Figure 1) (3). Thus, the physical removal from the conformation observed in the B-form helix explains how the cytosine C5 position becomes accessible to the enzyme. However, it is still unclear how the methylase extrudes the base from the double helix, either by actively inducing base flipping or by more passively capturing a transient conformation where the base is at least partially extended outside of the helix.
10.1021/bi7020469 CCC: $40.75 2008 American Chemical Society Published on Web 06/26/2008
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Earlier work has found that mutation of the target C to A, T, G, U, or m5C leads to the formation of stable DNA-protein complexes (4) and suggests that these complexes have flipped-out bases. Crystal structures demonstrated that when the target C base is mutated so that G:A, G:U, and G:AP (where AP is an abasic site) mismatch pairs are formed, the mismatched adenine and uracil are all flipped out into the enzyme active site pocket as in the cognate sequence (5). In the case of a G:AP mismatch, conformational changes occur in the target abasic nucleotide similar to those that effect base flipping, yet deleting any nucleotide in the recognition site from a position other than the target site leads to a loss of interaction with the methylase (4, 6). Thus, the identity of the extruded nucleotide is not important, yet base flipping is sequence specific. These observations suggested that the interaction between M.HhaI and the target DNA is not characterized solely by specific interactions between protein residues and specific positions on the target base but that recognition also requires interactions with the nonmethylated base pairs when the target base is correctly positioned (4). We have previously hypothesized that motions specific to the HhaI target sequence facilitate base extrusion by this enzyme, thereby making base flipping energetically feasible. Consistent with this hypothesis, solid-state NMR work has shown considerable sequence-dependent dynamics, on a microsecond to nanosecond time scale, across the four nucleotides recognized by the methylase (7). 19F NMR has been used in the past to characterize the solution-state motions within the cytosine that is the target for methylation, but these studies were limited because a single probe of dynamics was used (8). To probe more extensively the motional characteristics of this DNA sequence in solution, we used 13C NMR relaxation to study base and sugar dynamics for the DNA target of HhaI methyltransferase. In this work, we report 13C T1, T1F, and heteronuclear NOE values for sugar and base nuclei and the analysis of the motional properties of the HhaI DNA based on the modelfree formalism (9–11). Surprisingly, we found significant differences in the relaxation properties of the furanose rings of cytosines that are distinct from those observed for the other bases and much slower motions for the bases within the HhaI recognition sequence. To investigate whether the motions we observed are specific to the HhaI target sequence, we compare our work to solution (11) and solid-state NMR studies of the EcoRI DNA target (12–15), and to solid-state NMR studies of a similar construct of the HhaI DNA target (7). In both sequences, the furanose rings of the deoxycytidines have increased mobility compared to the other nucleotides, although the HhaI methylation site remains particularly flexible. In contrast, the slower (1-100 µs) motions observed for bases within the HhaI target sequence are not evident in the EcoRI target DNA (16). EXPERIMENTAL PROCEDURES DNA Preparation. A uniformly 13C- and 15N-labeled nonpalindromic dsDNA dodecamer with the sequence [5′(dGGTAGCGCTATT)-3′]-[5′-(dAATAGCGCTACC)-3′] (Figure 2) was purchased from BioQuantis. Each strand of DNA was dissolved and dialyzed into NMR buffer [10 mM potassium phosphate buffer, 50 mM KCl, and 0.1 mM EDTA
Shajani and Varani
FIGURE 2: Sequence of the HhaI target DNA. The recognition sequence is colored blue with the methylation site underlined.
(pH 6.0)]. After lyophilization, each strand was redissolved in 99.9% D2O, lyophilized again, and finally dissolved in 99.99% D2O, to remove any residual water. The two strands were combined, heated to 90 °C, and snap-cooled. The final sample concentration was 1.3 mM dsDNA. To demonstrate that the desired duplex was formed with a 1:1 stoichiometry, 13C HSQC experiments were performed first on the single-stranded DNAs. Each single strand yielded 24 resonances for the C6/C8 region at 25 °C, suggesting the formation of multiple conformations, probably a duplex in equilibrium with the single strands for these partially complementary sequences. However, at 45 °C, only 12 resonances could be seen in the C6/C8 region for each of the individual strands, suggesting that a single conformer could easily be formed by thermal denaturation. Indeed, when the two strands were annealed at 25 °C, 16 resonances were found, as expected for the desired duplex with no alternate conformers. Spectral Assignments. Proton and carbon assignments for the double-stranded DNA were obtained from the analysis of two-dimensional (2D) NOESY and three-dimensional (3D) 13 C NOESY-HSQC spectra recorded in 99.9% D2O at 25 °C using a Bruker Avance-500 spectrometer. Spectral assignments were based on standard NMR methods (17). A 2D NOESY spectrum collected with a mixing time of 200 ms was analyzed to generate initial proton assignments. The 3D 13C NOESY-HSQC experiment (mixing time of 120 ms) was then used to assign the carbon resonances. NMR Relaxation Experiments. T1, T1F, and heteronuclear NOE values were recorded as a series of 2D NMR spectra, in which the relaxation delay τ was parametrically increased. All experiments were performed in the constant-time mode. Data collection was performed on a Bruker Avance-500 instrument on the same 1.3 mM sample of DNA in 99.9% D2O at 25 °C. Data were collected on a TXI triple-resonance HCN probe. 15N or 31P decoupling was not introduced to reduce sample heating. The constant time delays were set to 7.15 and 12.5 ms when base and sugar relaxation properties were recorded, respectively, corresponding to 13C-13C coupling constants of approximately 70 Hz (bases) and 40 Hz (sugars) (18). The relaxation delay between scans was set to 1.9 s. Spectra were recorded with 86 complex points (base resonances) and 128 complex points (sugars) in the indirect dimension. The total collection time for each set of experiments was ∼24 h for either T1 or T1F measurements. Selective excitation was accomplished by application of 13 C 180° IBURP-shaped pulses during the first INEPT transfer (11, 19). An inversion bandwidth of 8 ppm was achieved by using pulses with a duration of 1 s at a 13C frequency of 125 MHz. The C6 and C5 resonances are separated by 40 ppm, enabling T1 to be easily measured for either nucleus without excitation of the other through the use of these selective pulses. Unlike in RNA, the ribose resonances are well dispersed in DNA and selective excita-
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tion is possible for C1′, C2′ (H′ and H′′), C3′, C4′, and C5′ (H′ and H′′) nuclei. Each spectral region required collection of an independent set of T1 values to ensure selectivity, whereas for T1F and NOE experiments, data for the C1′/C4′ and C3′/C5′ regions could be collected together. 13C T1 relaxation data were collected for the C8 resonances using nonselective pulses by setting the carrier frequency to 141 ppm. Using selective excitation pulses, 13C T1 relaxation data were collected for the C6 region in the bases (at 141 ppm), C1′ (carrier set to 91.5 ppm), C2′ (carrier set to 43 ppm), C3′ (carrier set to 60 ppm), and C5′/C5′′ (carrier set to 69 ppm) in the sugar. For T1F and NOE experiments, relaxation data were collected for C8 and C6 resonances (carrier set to 141 ppm), C1′ (carrier at 91.5 ppm), C2′ (T1F only, carrier set to 43 ppm), and C3′/C5′/C5′′ (carrier set to 69 ppm). T1F experiments were executed at a spin-lock field of 2 kHz. Delays of 5, 20, 40, 80, 120, 160, 240, 320, 500, 600, and 760 ms and 8, 12, 20, 40, 60, 80, and 96 ms were used for T1 and T1F experiments, respectively. One of the experimental points at random was repeated in a triplicate manner for each run to test the reproducibility of the measurements. For heteronuclear NOE measurements, a pair of spectra were recorded, one with initial proton saturation and one without. Proton saturation was achieved by the application of a train of 120° 1H pulses applied each 5 ms. Spectra recorded with proton saturation utilized a relaxation delay of 2.5 s followed by a 2.5 s period of saturation. Spectra recorded in the absence of saturation employed a total recycle delay of 5 s. Heteronuclear NOE spectra were collected at 11.7 T for all the base and sugar resonances. Power Dependence of T1F. Data reporting on the power dependence of T1F were collected and analyzed for C8-H8 and C6-H6 resonances at 11.7 T as a series of T1F experiments at various spin-lock field strengths (1.2, 2.6, 3.6, and 6.6 kHz). These values were set by modifying the power level of the spin-lock pulse. Data at each power level were collected and analyzed independently with the same settings and delays as described above for collection of T1F data. For the higher spin-lock fields (3.6 and 6.6 kHz), the longest relaxation delay was set to 28 ms to avoid sample heating. Data Analysis. All data were processed as described previously (20, 21). Briefly, T1 and T1F were determined from the decay curves using the standard equation I(τ) ) Io exp(-τ ⁄ T1 ⁄ 1F)
(1)
where Io is the initial peak intensity and τ is the delay time. The errors for the rate constants were estimated by Sparky and reflect the likely errors of the best fit T1/1F from the T1/1F values obtained for a perfect exponential decay; error estimates are reported in the Results. The heteronuclear NOE values were calculated from the ratio of the intensity of saturated to unsaturated spectra. Uncertainties in these measurements were estimated from the base plane noise in the 2D 1H-13C HSQC spectra recorded with and without proton saturation, as described previously (22). Relaxation times were derived using all time delays mentioned above; reassuringly, the results did not change by more than 2% when only the first six delay points were fitted, indicating that relaxation is monoexponential. Thus, cross-talk between neighboring 13C nuclei is effectively minimized by the use of selective excitation pulses (11).
FIGURE 3: Relaxation times for C3′-H3′ spins: (a) 13C T1, (b) 13C T1F, and (c) heteronuclear NOE. Residues that had noisy relaxation curves or that were partially or completely overlapped were excluded from the analysis.
Analysis of Relaxation Data with the Modelfree Approximation. Quantitative analysis of the base (C8-H8 and C6-H6 resonances) and sugar C1′-H1′ and C3′-H3′ relaxation data was conducted using the model-free approximation (9, 10) with the program ModelFree 4.15 (23) in a manner analogous to previous studies of RNA (20, 24). Model selection was conducted according to previously reported procedures (23, 25). Due to additional relaxation mechanisms between the strong coupled H2′ and H2′′ nuclei, the C2′-H2′ resonances were not analyzed quantitatively. The rotational correlation time for the entire molecule was found to be 4 ns for all models with a Dratio of 2.0 and C-H bond lengths of 1.09 and 1.104 Å for the ribose and bases, respectively. For this analysis, we used the values of chemical shift anisotropies (CSAs) that were recently reported for doublestranded B-form DNA (144 ppm for dA-C8, 133 ppm for dG-C8, 168 ppm for dT-C6, and 186 ppm for dC-C6) for the base resonances (26). These CSA values also agree well with those determined by solid-state NMR on DNA mononucleosides (27). The CSA used for ribose resonances was 30 ppm, as determined by Bryce and co-workers (28) and Stueber and Grant (27). For the C1′-H1′ resonances, three resonances were fit to model 2 (T9, T11, and A14), eight resonances to model 5 (G1, T3, A4, G7, C8, A10, T12, and A13), and the remaining five resonances (G2, G5, C6, C23, and C24) to model 4. For the C3′-H3′ resonances, one resonance was fit to model 2 (A13) and two resonances were fit to model 5 (T12 and C24); the remaining residues were fit to model 4. However, it was not possible to fit the base resonances, likely because many of them exhibited exchange (see below). RESULTS Data Collection and Analysis. 13C T1, T1F, and heteronuclear NOE values were recorded as described in Experimental Procedures. All relaxation decay curves could be fit very well by single exponentials, without any evidence of multiexponential behavior, suggesting that cross-talk between neighboring spins was minimized by the use of selective excitation pulses. The observed relaxation times obtained from the analysis of these decay curves and the heteronuclear NOEs measured at 11.7 T are shown in Figures 3–6. This
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Shajani and Varani Table 1: Average Relaxation Times and NOE Ratios for Ribose Resonances (excluding 5′ and 3′ terminal residues) C1′-H1′ C2′-H2′ C2′-H2′′ C3′-H3′ C4′-H4′
T1 (ms)
T1F (ms)
NOE ratio
413 (0.8%) 151 (9.7%) 233 (6%) 424 (1.0%) 410 (0.8%)
59 (0.6%) 42 (2.4%) 42 (2.4%) 59 (1.0%) 57 (0.8%)
1.36 (2%) 1.57 (2%) 1.55 (2%) 1.40 (2%) 1.38 (2%)
Table 2: Average Relaxation Times and NOE Ratios for Base Resonances (excluding 5′ and 3′ terminal residues)
FIGURE 4: Relaxation times for C2′-H2′′ spins: (a) 13C T1, (b) 13C T1F, and (c) heteronuclear NOE. Residues that had noisy relaxation curves or that were partially or completely overlapped were excluded from the analysis.
FIGURE 5: Relaxation times for C1′-H1′ spins: (a) 13C T1, (b) 13C T1F, and (c) heteronuclear NOE. Residues that had noisy relaxation curves or that were partially or completely overlapped were excluded from the analysis.
FIGURE 6: Relaxation times for C6-H6 and C8-H8 spins: (a) 13C T1, (b) 13C T1F, and (c) heteronuclear NOE. Residues that had noisy relaxation curves or that were partially or completely overlapped were excluded from the analysis.
field was chosen to minimize artifacts in the analysis arising from the noncollinearity of the CSA with the C-H bond and the large CSA of base nuclei (20, 29). Chemical shift assignments were obtained using standard methods based on the analysis of 2D NOESY and 3D 13Cedited NOESY-HSQC spectra; assignments have been deposited into the BioMagResBank. Some nuclei were excluded from the analysis of the relaxation data because
A: C8-H8 G: C8-H8 T: C6-H6 C: C6-H6
T1 (ms)
T1F (ms)
NOE ratio
392 (0.9%) 379 (0.9%) 341 (1.1%) 322 (1.1%)
35 (0.9%) 28 (0.9%) 34 (0.7%) 29 (0.7%)
1.22 (1.8%) 1.19 (1.8%) 1.18 (1.8%) 1.17 (1.8%)
the corresponding resonances were overlapped. However, the 13 C relaxation properties for six of eight purine C8, all of the pyrimidine C6 (eight of eight), and all ribose C1′ (16 of 16) resonances could be measured and analyzed reliably. The 13 C relaxation properties of 13 of 16 C2′-H2′ and C2′-H2′′ peaks and 14 of 16 C3′-H3′ peaks could also be measured and analyzed. Unfortunately, most of the resonances in the C5′-H5′ and C5-H5′′ region were overlapped and could not be resolved sufficiently well to warrant analysis. QualitatiVe Analysis of the 13C Relaxation Data. The average relaxation rates are reported first, since they provide a reference point for establishing which residues deviate from the behavior expected for a perfectly rigid duplex (Tables 1 and 2). The data demonstrate the presence of noticeably less restricted motions at the ends of the sequence, which is expected as the terminal bases fray in solution. The T1F and NOE values are substantially larger than the average observed in the rest of the helix, suggesting the presence of extensive motions on the picosecond to nanosecond time scale. Thus, the 5′ and 3′ terminal residues were excluded from the averages listed in Tables 1 and 2 because of anomalous motional properties due to well-known end effects (base fraying). More specifically, the C3′-H3′ resonances of the terminal G1 residue and C24 residues have T1F values that are 55 and 210% (90 and 135 ms, respectively) greater than average and NOE ratios that are 34 and 23% (1.80 and 1.85, respectively) greater than average, respectively. The terminal T12 and A13 residues have T1F values that are 250 and 57% (141 and 90 ms, respectively) greater than average and NOE values that are 27 and 14% (1.80 and 1.64, respectively) greater than average, respectively. Surprisingly, all the deoxycytidines (terminal residues are not considered in this analysis) have T1F values that are 8-12% and NOE values that are 5-12% larger compared to those of the deoxyadenosines, deoxyguanosines, and deoxythymidines, implying that the riboses of the deoxycytidines are somewhat more flexible on a picosecond to nanosecond time scale compared to the other nucleotides. The C2′-H2′′ resonance of C24 has a T1F value that is 66% (75 ms) greater than average and a NOE ratio that is 20% (1.86) greater than average. The T12 residue and A13 residue have T1F values that are 98% (83 ms) and 31% (54 ms) greater than average and NOE ratios that are 20% (1.86) and 12% (1.73) greater than average, respectively.
Dynamics of HhaI Target DNA As with the C3′-H3′ resonances, the deoxycytidines (not considering terminal residues) have slightly larger T1F values (7%) than the deoxyadenosines, deoxyguanosines, and deoxythymidines. For the C1′-H1′ resonances as well, the 5′ G-C and 3′ A-T pairs have larger T1F and NOE values, but to a lesser extent compared to the C2′-H2′′ and C3′-H3′ resonances. This difference probably reflects the larger effect on C2′ and C3′ of the transition of the furanose ring between C2′- and C3′-endo conformers that is often observed for nucleotides at the 5′ and, especially, 3′ end of helices. Thus, the G1 residue has a T1F value that is 17% (68 ms) greater than average and a NOE ratio that is 25% (1.69) greater than average. The T12 and A13 residues have T1F values that are 53% (95 ms) and 43% (79 ms) greater than average and NOE values that are 18% (1.62) and 20% (1.60) larger, respectively. Unexpectedly, the average T1F values (again not considering the terminal residues) for both (deoxycytidine and deoxythymidine) are 12% higher than those of the purines. The increased flexibility of the C1′-H1′ deoxycytidines reflected in this anomalous relaxation rate is consistent with what is observed for other ribose carbons. Overall, these results suggest that the entire furanose ring of cytosines experiences increased motional freedom compared to the three other nucleotides. As observed for the sugar rings, the base resonances of the 5′ and 3′ terminal residues also display anomalous dynamics, suggesting the presence of extensive motions on the picosecond to nanosecond time scale. The C6-H6 base resonance of the T12 residue has T1F and NOE values that are 61 and 20% (55 ms and 1.42, respectively) greater than average, respectively. The C8-H8 base resonance for the A13 residue has T1F and NOE values that are 34 and 11% (47 ms and 1.35, respectively) greater than average, respectively. The C8 base resonance for the G1 residues has a NOE value that is 20% (1.36) greater than average. The C6 resonance for the C24 residue has T1F and NOE values that are 20 and 190% (35 ms and 1.39, respectively) greater than average, respectively. Analysis of the average transverse relaxation times for each type of base shows unexpected differences which are well beyond any experimental uncertainty (Table 2). Given the magnitude of the CSA, we would have predicted that the purine (G and A) bases would have very similar relaxation times if the DNA helix behaved as a perfectly rigid molecule without any significant internal motion, and the pyrimidine bases (T and C) would also have similar relaxation times. Instead, we observe that the G and C bases have average T1F relaxation times that are 20% smaller than those of the A and T bases. However, for each particular type of base, there are no large differences across the sequence among dA, dC, dG, and dT (excluding the terminal residues); i.e., C6, C8, and C23 all have similar values, with one significant exception. The A4 base residue has a T1F value (30 ms) lower than those of the other two A’s (37 ms); interestingly, A4 is adjacent to the HhaI recognition sequence GCGC. These differences in relaxation times cannot be attributed to the CSA or to anisotropic tumbling (see below) and reflect real differences in the internal dynamics of this DNA sequence. Interestingly, the longitudinal relaxation times do not follow this trend but rather behave as would be predicted from the relative values of the CSA, with the pyrimidine
Biochemistry, Vol. 47, No. 29, 2008 7621 bases (which have the largest CSA) exhibiting the smallest T1 values. This result strongly suggests that the differences in relaxation times between different nucleotides can be attributed to motions on time scales (microseconds to milliseconds) which affect only the transverse but not the longitudinal relaxation times. Power Dependence of T1F for Base Resonances. The qualitative analysis of the previous paragraph attributed the reduction of dG and dC (as well as A4) base transverse relaxation times to motions on microsecond to millisecond time scales. To test this hypothesis, we measured the power dependence of T1F. Resonances that exhibit conformational exchange on the time scale corresponding to the power of the spin-lock pulses will display a decrease in the relaxation rate as the spin-lock field strength increases. Consistent with our interpretation of the primary relaxation times, we observe a slight power dependence of T1F for all of the deoxycytidine resonances (Figure 7). This result demonstrates that the dC base residues are involved in conformational exchange processes, and the shape of the power dependence curves demonstrates that the rates of these processes are close to the microsecond time scale. Furthermore, G5 exhibited a strong initial dependence of relaxation rate on spin-lock power (Figure 7e) before leveling out, suggesting that motions occur at a rate that is at the edge of what can be probed with available power levels (e.g., 1-10 µs). These studies provide no evidence of conformational exchange processes for A4, G1, or G7, suggesting that the anomalous behavior of A4 and G7 may reflect motions on time scales that are too fast to be probed with current relaxation dispersion techniques. ModelFree Analysis of the 13C Relaxation Data. Quantitative analysis of the relaxation data using the model-free approximation was conducted with ModelFree 4.15 as described in Experimental Procedures. Data were best fit with an axially symmetric diffusion model, with a diffusion anisotropy of 2.0 and an overall correlation time (τm) of 4.0 ns. Appropriately, τm and the diffusion anisotropy were determined by excluding 5′ and 3′ terminal bases (because of an increased level of internal motion); this property was also calculated twice independently for C1′ and C3′ resonances with similar results to further validate the results. Previous studies on DNAs similar in size reported correlation times ranging from 3.3 to 5 ns (16, 30). A summary of the parameters that best describe the dynamics of each residue is given in Figure 8 and Table 3; errors for the order parameters are ∼3% as determined with ModelFree 4.15. Residues that were fit best with model 4 required only low levels of Rex (
